Utilization of Piper betle L. Extract for Inactivating Foodborne Bacterial Biofilms on Pitted and Smooth Stainless Steel Surfaces

Biofilms are a significant concern in the food industry. The utilization of plant-derived compounds to inactivate biofilms on food contact surfaces has not been widely reported. Also, the increasing negative perception of consumers against synthetic sanitizers has encouraged the hunt for natural compounds as alternatives. Therefore, in this study we evaluated the antimicrobial activities of ethanol extracts, acetone extracts, and essential oils (EOs) of seven culinary herbs against Salmonella enterica serotype Typhimurium and Listeria innocua using the broth microdilution assay. Among all tested extracts and EOs, the ethanol extract of Piper betle L. exhibited the most efficient antimicrobial activities. To evaluate the biofilm inactivation effect, S. Typhimurium and L. innocua biofilms on pitted and smooth stainless steel (SS) coupons were exposed to P. betle ethanol extract (12.5 mg/ml), sodium hypochlorite (NaClO; 200 ppm), hydrogen peroxide (HP; 1100 ppm), and benzalkonium chloride (BKC; 400 ppm) for 15 min. Results showed that, for the untreated controls, higher sessile cell counts were observed on pitted SS versus smooth SS coupons. Overall, biofilm inactivation efficacies of the tested sanitizers followed the trend of P. betle extract ≥ BKC > NaClO > HP. The surface condition of SS did not affect the biofilm inactivation effect of each tested sanitizer. The contact angle results revealed P. betle ethanol extract could increase the surface wettability of SS coupons. This research suggests P. betle extract might be utilized as an alternative sanitizer in food processing facilities.


Introduction
Foodborne illnesses have been of paramount concern to the food industry. The US Centers for Disease Control and Prevention estimates approximately 48 million sick patients, 128,000 hospitalizations, and 3,000 deaths from foodborne illnesses per year [1]. According to the Thailand Bureau of Epidemiology, more than 100,000 annual cases of foodborne illness have been reported over the past decade [2]. Food can become contaminated at any point from farm to table as well as during manufacturing, transportation, and even in the consumer's kitchen; consuming contaminated food will eventually result in foodborne illnesses. Ineffective cleaning and sanitizing of food contact surfaces (FCSs) in food processing plants can promote soil buildup and, in the presence of water, contribute to the formation of bacterial biofilms which may harbor pathogens [3]. Biofilms are spatially structured communities of microorganisms attached to and growing on surfaces embedded in extracellular polymeric substances (EPSs) consisting of polysaccharides, proteins, nucleic acids, and lipids [3,4]. Salmonella spp. and Listeria monocytogenes are bacterial pathogens that can form biofilms on FCSs and have been associated with many foodborne illnesses [5][6][7]. Once formed on an FCS, biofilms can become more resistant to cleaning and sanitizing agents, rendering it more difficult to remove [3,8]. It has been reported that commonly used sanitizers (e.g., sodium hypochlorite, sodium hydroxide, hydrogen peroxide, and peroxyacetic acid) failed to eradicate biofilms on FCSs [9,10]. Some commonly used sanitizing agents may also produce harmful by-products (e.g., chloramine, trihalomethane, haloacetic acid, and other potentially carcinogenic compounds), cause microbial resistance [11], and corrode FCSs [12,13].
Biofilms are a significant concern in the food industry. The utilization of plant-derived compounds to inactivate biofilms on food contact surfaces has not been widely reported. Also, the increasing negative perception of consumers against synthetic sanitizers has encouraged the hunt for natural compounds as alternatives. Therefore, in this study we evaluated the antimicrobial activities of ethanol extracts, acetone extracts, and essential oils (EOs) of seven culinary herbs against Salmonella enterica serotype Typhimurium and Listeria innocua using the broth microdilution assay. Among all tested extracts and EOs, the ethanol extract of Piper betle L. exhibited the most efficient antimicrobial activities. To evaluate the biofilm inactivation effect, S. Typhimurium and L. innocua biofilms on pitted and smooth stainless steel (SS) coupons were exposed to P. betle ethanol extract (12.5 mg/ml), sodium hypochlorite (NaClO; 200 ppm), hydrogen peroxide (HP; 1100 ppm), and benzalkonium chloride (BKC; 400 ppm) for 15 min. Results showed that, for the untreated controls, higher sessile cell counts were observed on pitted SS versus smooth SS coupons. Overall, biofilm inactivation efficacies of the tested sanitizers followed the trend of P. betle extract ≥ BKC > NaClO > HP. The surface condition of SS did not affect the biofilm inactivation effect of each tested sanitizer. The contact angle results revealed P. betle ethanol extract could increase the surface wettability of SS coupons. This research suggests P. betle extract might be utilized as an alternative sanitizer in food processing facilities.

Determination of Minimum Inhibitory and Minimum Bactericidal Concentrations
The broth microdilution assay was performed to determine the minimum inhibitory concentration (MIC) and the minimum bactericidal concentration (MBC) values of plant extracts and EOs as described by Damrongsaktrakul et al. [23]. TSB and TSBYE were used for culturing S. Typhimurium and L. innocua, respectively. The plant extracts and EOs (100 μl) were serially 10-fold diluted in 20% DMSO in a 96-well plate to obtain initial concentrations ranging from 0.78 to 400 mg/ml. Then, 100 μl of S. Typhimurium and L. innocua in 2X TSB and TSBYE were added to the wells to obtain a concentration of approximately 5 log CFU/ml; the final concentrations of the plant extracts and EOs were 0.39 to 200 mg/ml in 10% DMSO. Negative growth controls (extracts and EOs in the test medium without bacteria) and positive growth controls (bacteria in the test medium + 10% DMSO without extracts and EOs) were also included. The plates were incubated at 37°C for 24 h. Then, 30 μl of 0.015% resazurin (Sigma Aldrich Co., USA) was added to the wells. After 30-min incubation, the plates were visually inspected. The lowest concentrations of the extracts and EOs that inhibited the growth of tested microorganisms (i.e., no red or pink color) were considered the MIC. To determine the MBC values, 100 μl of the test solutions from inhibitory wells were spread on TSA plates. The plates were incubated at 37°C for 24 h. The lowest concentrations of extracts and EOs yielding ≥ 3.0 log CFU/ml reduction were deemed the MBC. The assay was completed in triplicate; each replicate was performed in duplicate (n = 6).

Determination of Total Phenolic Content of P. betle Extract
To determine the total phenolic content of the P. betle ethanol extract, a Folin-Ciocalteu assay was performed as described by Damrongsaktrakul et al. [23] with minor modifications. The standard gallic acid solutions were prepared using 95% ethanol to obtain concentrations of 0 to 400 mg/l. The P. betle ethanol extract was dissolved in 95% ethanol to obtain a concentration of 500 mg/ml. Then, the standard gallic acid solutions and the P. betle extract were individually mixed with 0.5 ml of distilled water and 125 μl of Folin-Ciocalteu reagent, respectively. The resulting solutions were allowed to incubate for 6 min. After that, 1.25 ml of 7% sodium carbonate solution was added, followed by distilled water to bring the final volume to 3 ml. The resulting solutions were then incubated for 90 min and measured at 760 nm using a spectrophotometer (BioTek, USA) with a reference blank. The total phenolic content was represented as milligrams of gallic acid equivalent in a gram of crude extract (mg GAE/g).

Preparation of Pitted and Smooth Stainless Steel Coupons
The type 304 SS coupons (1 cm × 2 cm) were kindly provided by AEC Industrial Services Co., Ltd. For the smooth surface condition, the SS coupons were polished using a fine grit sandpaper and rinsed with distilled water. The pitted surface condition was prepared using an electrochemical technique, according to Asaduzzaman et al. [24], with modification. The polished SS coupons were coated with plastic tape leaving a 1 cm × 1 cm area for electrolyte exposure. The coated SS coupons were then submerged in 50 ml of 3.5% NaCl. The DC power supply YG3005D (Yugo, Thailand) was used to provide an electrical potential (5 V) for 5 min to induce corrosion in order to generate pits on the SS coupons at room temperature. The pitted and smooth SS coupons were then cut into a size of 1 cm × 1 cm and submerged in acetone for 10 min. To sterilize them, the SS coupons were soaked in 95% ethanol for 15 min. Then, the coupons were autoclaved at 121°C for 15 min and dried in a hot air oven at 60°C for 24 h.

Pit Measurement
Pits were observed under a Carl Zeiss Axiovert 40 MAT optical microscope (Carl Zeiss, Germany) equipped with a Dino-Eye Edge AM7025X digital camera (Dino-Lite, Taiwan), and the density of pits was expressed as numbers of pits/mm 2 . The pit diameter was determined using DinoCapture 2.0 microscope imaging software (Dino-Lite). Pit depth was determined by measuring the depth of pit cross-sections under the optical microscope using DinoCapture 2.0 microscope imaging software.

Biofilm Formation on Stainless Steel Coupons
A cocktail inoculum of S. Typhimurium and L. innocua was prepared as described by Yegin et al. [25] with minor modifications. S. Typhimurium and L. innocua were centrifuged at 10,000 ×g at 22°C for 15 min, and the supernatants were discarded. The resulting pellets were washed in 10 ml of phosphate-buffered saline (PBS; HiMedia) and centrifuged for 15 min. Cell washing and centrifugation were completed twice. Then, S. Typhimurium and L. innocua pellets were resuspended in 0.1% peptone water (PW; HiMedia) and mixed. To form biofilm on SS coupons, the sterile coupons were placed in conical tubes containing 10 ml of TSB. Then, the cocktail inoculum was added to the tubes to achieve a target concentration of approximately 7 log CFU/ml. The tubes were incubated at 30°C for 48 h to allow biofilm formation.

Sanitization Treatment
Following 48 h of incubation, the SS coupons were withdrawn from the conical tubes and washed three times using PBS to remove loosely attached cells. To prepare sanitizers, P. betle extract was dissolved in 10% DMSO to obtain the final concentration of 12.5 mg/ml. Sodium hypochlorite (NaClO; Loba Chemie, India), hydrogen peroxide (HP; QRec, New Zealand) and benzalkonium chloride (BKC, Hong Huat, Thailand) were dissolved in sterile distilled water to obtain the final concentrations of 200 ppm, 1,100 ppm, and 400 ppm, respectively. For sanitization treatment, the coupons were individually submerged in conical tubes containing 4 ml of P. betle ethanol extract, NaClO (pH 7), HP, and BKC for 15 min. The untreated control samples were included to assess sessile cell numbers of S. Typhimurium and L. innocua on the SS coupons without sanitization. After sanitizing treatment, the coupons were washed twice with PBS to eliminate sanitizer residues and loosely attached cells. The coupons were placed in conical tubes bearing 10 ml of 0.1% PW and sonicated at 50 Hz for 1 min (Sonics & Materials, Inc., USA). The dislodged biofilm was serially diluted in 9 ml of 0.1% PW and spread on XLD and Oxford agar plates. The resulting plates were incubated at 37°C for 24 h, and the microbial colonies were then enumerated. The limit of detection (LOD) of the assay was 0.7 log CFU/cm 2 . The assay was performed in triplicate, with two independent samples being tested per replicate (n = 6).

Scanning Electron Microscopy Analysis
The untreated control and sanitizer-treated biofilm samples on pitted and smooth SS coupons were subjected to scanning electron microscopy (SEM) analysis. The coupons were washed with PBS and distilled water. The sample fixation was done by submerging the samples in 4% glutaraldehyde (Fisher Scientific) at 4°C for 2 h. Then, the coupons were washed with distilled water and gradually dehydrated in an ethanol series of 25, 50, 75, 95, and 100%, respectively. Following dehydration, samples were subjected to PdAu sputter coating and then observed under SEM (Quanta 450, FEI, USA) [23].

Contact Angle Measurement
The contact angle measurement was performed as described by Zezzi et al. [26] with modifications. The pitted and smooth SS coupons were submerged in 4 ml of P. betle for 15 min and then washed with distilled water. To determine the effect of 10% DMSO (solvent for P. betle extract) on the contact angles, the SS samples were also treated with 4 ml of 10% DMSO for 15 min and then rinsed with distilled water. For the control samples, the SS coupons were immersed in distilled water for 15 min. All SS samples were then dried in a desiccator for 24 h. A sessile drop technique was used to assess the contact angle by dropping 2 μl of distilled water on the SS samples. The contact angle was measured after 10 s deposition to allow the water droplet to stabilize. An optical contact angle and interface tension meter (Model SL150, USA Kino, China) was used for the contact angle measurement.

Statistical Analysis
Two-way analysis of variance (ANOVA) at α = 0.05 was used to determine mean differences in the populations of the tested pathogens as a function of the main effects of sanitizing treatment, surface condition of SS, and their interaction via PASW Statistics software version 17.02 (SPSS Inc., USA). The contact angle results were analyzed using one-way ANOVA. Mean separation of the pathogens and the contact angle data were performed using Fisher's least significant difference (LSD) procedure. For the purpose of statistical analysis, 0.4 log CFU/cm 2 was assigned to the non-detectable level of the tested pathogens.

Extraction Yields
The plant extraction yields are presented in Table S1. The yields from ethanol and acetone extraction ranged from 0.94 to 6.14% w/w and 0.78 to 5.84% w/w, respectively. Among all tested plant extracts, the highest yield was observed with the ethanol extract of P. betle (6.14% w/w). The plant parts and the type of solvents played a crucial role in the crude extract yields. Different types of organic solvents vary in polarity and capacity to permeate plant cell walls. Hence, the targeted products were distinctly dissolved and separated from the structural components [27]. Ethanol extract yields of P. betle have been reported in previous studies [28,29]. Boontha et al. [28] reported 1.4% w/w yield, while Pin et al. [29] demonstrated approximately 10% yield for the ethanol extract of P. betle. Yield variation could be due to phytochemicals in plants, the extraction method, the extraction temperature, and the ratio of solvent to solid [29,30]. Table 1 shows the MICs and MBCs of ethanol extracts, acetone extracts, and essential oils (EOs) of the tested plants against S. Typhimurium and L. innocua. MIC and MBC values against S. Typhimurium ranged from 3.125 to 200 mg/ml and 12.5 to >200 mg/ml, respectively. Overall, the acetone extract of C. citratus was the least inhibitory and bactericidal to S. Typhimurium. For L. innocua, MIC and MBC values varied from 3.125 to 100 and 12.5 to 200 mg/ml, respectively. For both tested microorganisms, the ethanol extract of P. betle displayed the lowest MICs and MBCs, and thus indicated the highest antimicrobial activity among the ethanol extracts, acetone extracts, and EOs of the tested plants. Growth of the tested pathogens was observed in the control wells, indicating that 10% DMSO showed no inhibitory effect against S. Typhimurium and L. innocua.

Minimum Inhibitory and Minimum Bactericidal Concentrations
Hydroxychavicol, eugenol, and gallic acid have been reported to be significant phytochemical components of P. betle leaf, among others [31]. Hydroxychavicol has two hydroxyl groups and is more soluble in the solvent with high polarity [29]. Nguyen et al. [31] reported that P. betle leaf extracts using polar solvents showed more efficient antimicrobial activities than that with a nonpolar solvent. High contents of hydroxychavicol and eugenol were also observed with ethanol extract of P. betle [31]. In our study, the ethanol extract of P. betle exhibited higher antimicrobial activity than the acetone extract. Ethanol has higher polarity than acetone [32]. It may have dissolved hydroxychavicol and other polar or semi-polar components more efficiently than acetone, thus resulting in the P. betle ethanol extract having a higher amount of phytochemical components and antimicrobial activity. Previous findings have shown that P. betle extract exerted no significant adverse effects on human cells [33], rats, and mice [34][35][36], suggesting the safety of P. betle extract for human use.

Total Phenolic Content of P. betle Extract
The total phenolic content of P. betle ethanol extract was 444 mg GAE/g. The result was within the range of the total phenolic content values reported by previous studies [27,31,37]. Factors affecting differences in the phenolic content across studies could be the extraction temperature, extraction method, age of plants, and conditions of the phenolic content assay.

Pit Characteristics
The optical microscope images of pits on SS surfaces formed by an electrochemical technique are demonstrated in Fig. 1. According to ASTM G46-94 (2018) [38], pits in this study appeared to have a "wide, shallow" shape ( Fig. 1B). It should be noted that the pit formation was conducted for 5 min; therefore, longer exposure time to an electrical potential may have resulted in changing the shape of pits. The average size of simulated pit mouths was 63.3 ± 27.3 μm in diameter. Several aspects, such as the inhomogeneity of the SS alloys during manufacture, could cause a broad range of pit sizes. This could result in different resistance to corrosion locally. Therefore, the pit size, which was the result of pitting corrosion, could be varied accordingly. The depth of the simulated pits was 25.7 ± 10.6 μm. The same reason was applied to the variation of the pit depth. In addition, the method for determining pit depth could also affect the measurement results, such as the selection of the pit for evaluation and the pit crosssectioning method used. Another aspect of determining the extent of the pitting is the pit density, which in this case was 41.6 ± 9.2 pits/mm 2 .

Biofilm Inactivation Effect of Tested Sanitizers
According to the MIC and MBC results, the ethanol extract of P. betle was selected for the biofilm inactivation assay compared with the commonly used sanitizers, i.e., NaClO, HP, and BKC. Fig. 2 shows the means of surviving S. Typhimurium and L. innocua obtained from the mixed-culture biofilms on the pitted and smooth SS coupons following sanitization treatment. For S. Typhimurium, the sessile cell population on the untreated, pitted SS control (6.6 ± 0.2 log CFU/cm 2 ) was significantly higher (p < 0.05) than those on the untreated, smooth SS control (6.0 ± 0.4 log CFU/cm 2 ) ( Fig. 2A). After sanitization treatment, the S. Typhimurium cell numbers were reduced to the range of below detectable level to 3.5 ± 0.7 log CFU/cm 2 . For L. innocua, a significant difference (p < 0.05) was  Fig. 2. Means of surviving S. Typhimurium (A) and L. innocua (B) obtained from the mixed-culture biofilms on the pitted and smooth stainless steel coupons following sanitization treatment. The sanitizers were 12.5 mg/ml P. betle extract (PB), 200 ppm sodium hypochlorite (NaClO; pH 7), also observed between the sessile cell numbers on the untreated, pitted SS control (6.4 ± 0.2 log CFU/cm 2 ) and the population on the untreated, smooth SS control (5.8 ± 0.4 log CFU/cm 2 ) (Fig. 2B). The sessile cell numbers of L. innocua ranged from below detectable level to 3.5 ± 0.3 log CFU/cm 2 after treatment with the tested sanitizers. For each tested pathogen, the sessile cell numbers did not differ when treated with the same sanitizer (p > 0.05) regardless of the SS surface conditions. Our preliminary data showed that 10% DMSO had no significant inactivation effect on biofilms of S. Typhimurium (Table S2) and L. innocua (Table S3).
Overall, the biofilm inactivation effect of the tested sanitizers followed the pattern P. betle extract  BKC > NaClO > HP. Piper betle L. is a medicinal plant that belongs to the Piperaceae family with an origin in Southeast Asia [39]. It has been reported to possess anticarcinogenic, antimutagenic, anti-inflammatory, antibiofilm, and antimicrobial activities [31,40]. However, the study on the biofilm inactivation efficiency of P. betle extract on FCSs is still lacking. In this study, P. betle extract exhibited the most significant biofilm inactivation effect among the tested sanitizers. The qualitative phytochemical analysis by Syahidah et al. [41] demonstrated the presence of phenols, flavonoids, alkaloids, tannins, saponins, glycosides, terpenoids, and steroids in the P. betle leaf extract. Hydroxychavicol and eugenol are the major phenolic compounds in P. betle leaves [31,42]. Hydroxychavicol could induce DNA damage and suppress cell division, resulting in bacterial cell death [43]. For eugenol, the hydroxyl group could interact with microbial proteins and disrupt the enzyme function, leading to non-specific membrane permeability and inhibition of ions and ATP transport [31]. NaClO, HP, and BKC are commonly used sanitizers in food processing facilities [44]. NaClO has strong oxidizing activity and can oxidize the sulfydryl groups on certain enzymes in the cell membrane or protoplasts, resulting in impairment of cellular protein activities. Also, it may induce irreversible decarboxylation reactions [45]. HP is a strong oxidant and has been reported to damage bacterial DNA, proteins, and cellular membranes [46]. BKC is a quaternary ammonium compound; it can perturb and disrupt the membrane bilayers by the alkyl chains and disrupt the charge distribution of the microbial membrane by the charged nitrogen [47]. Robbins et al. [46] demonstrated 100 ppm chlorine decreased the biofilm of L. monocytogenes on stainless steel chip by 6.49 log CFU/chip after 10 min of exposure. Damrongsaktrakul et al. [23] showed 3.37 log CFU/cm 2 reduction of S. Typhimurium on SS samples after 30 min treatment with NaClO. The study by Kostaki et al. [48] revealed that 6 min exposure to 50 ppm BKC produced < 4 log CFU/cm 2 and < 3 log CFU/cm 2 reduction of the biofilms of S. enterica and L. monocytogenes (monospecies and dual species), respectively. In our study, NaClO and BKC reduced the biofilms of S. Typhimurium and L. innocua on pitted and smooth SS samples by > 5 log CFU/cm 2 . Nevertheless, NaClO failed to decrease biofilms of the tested pathogens to below the detectable level, while BKC was able to do so for L. innocua biofilms. The variation in sanitizer efficacy across studies might be attributable to the concentration of sanitizers, contact time, temperature, biofilm age, growth condition, and microbial type. For HP, reduction of < 4 log CFU/cm 2 of the tested pathogens was obtained in our study. S. Typhimurium and L. innocua are catalase-positive microorganisms [49,50]. Therefore, HP might be degraded by the tested pathogens, making it less efficient to inactivate biofilms than other tested sanitizers.
Pits and crevices could facilitate bacterial adhesion and biofilm formation on SS surfaces [9,16]. In the present study, the sessile cell numbers in the biofilms of S. Typhimurium and L. innocua on pitted SS were significantly greater than those on smooth SS samples. However, the sessile cell numbers of each tested pathogen on smooth versus pitted SS surfaces were not significantly different when treated with the same sanitizer. This may have been due to the large pit size on SS surfaces, which allowed sanitizer access to the pits. In agreement with our study, Kim et al. [51] reported higher counts of Staphylococcus aureus biofilm on scratched SS surfaces vs. smooth SS surfaces. No significant difference in chlorine (200 ppm) efficacy against biofilms on smooth and scratched SS surfaces was observed. In another study, the efficacy of 200 ppm chlorine dioxide, 70% alcohol, and 200 ppm quaternary ammonium compound against the biofilm of Bacillus cereus did not differ on smooth versus scratched SS surfaces after 10 min sanitization [52]. However, the sizes of scratches or crevices were not reported in these studies.

Scanning Electron Microscope Imaging
The SEM images of the mixed culture of S. Typhimurium and L. innocua biofilms on pitted and smooth stainless steel coupons after treatment with sanitizers are represented in Fig. 3. The untreated control samples on both SS surface conditions (Figs. 3A, 3a) showed intact, rod-shaped cell morphology with EPS. In addition, the sessile cells of the mixed culture were also observed in the pit of the SS coupon. The samples treated with P. betle extract (Figs. 3B, 3b) and BKC (Figs. 3E, 3e) showed a great reduction in the sessile cell numbers on both SS surface conditions. However, less reduction in the sessile cell numbers was observed with samples exposed to NaClO (Figs. 3C, 3c) and HP (Figs. 3D, 3d). A thick EPS layer was still present in the sample treated with HP (Fig. 3d).
In agreement with the plate count results, the SEM images showed greater amounts of sessile cells of the mixedculture biofilms on pitted versus smooth SS coupons. This indicated pits/crevices could allow for microbial attachment and biofilm formation. In line with our findings, the SEM images by Kim et al. [51] revealed more sessile cells of S. aureus biofilm on scratched SS surfaces compared to smooth surfaces. In our study, P. betle extract showed the greatest biofilm removal on both SS surface conditions, suggesting that P. betle extract may be employed as an alternative to the commonly used sanitizers in food processing facilities. For other tested sanitizers, lower numbers of sessile cells were observed after sanitization, indicating that these sanitizers could partially remove biofilms of S. Typhimurium and L. innocua. In addition to inactivating microorganisms, chlorine is known to remove EPS from surfaces [53,54]. Agreeing with our study, SEM images from previous research showed that commonly used sanitizers failed to completely eradicate microbial biofilms on SS surfaces [23,55]. It is worth noting that our study employed the submersion method for sanitization. Therefore, the sanitization efficacy may be improved by the combination of chemical, biological, and physical treatment [56].

Contact Angles of Stainless Steel Samples
To determine how P. betle extract affected the wettability of the SS samples, the pitted and smooth SS samples treated with P. betle extract were subjected to contact angle measurements ( Table 2). For either surface condition, 10% DMSO showed no significant effect (p > 0.05) on the contact angle values of the SS coupons versus controls. The contact angle values of the SS samples treated with P. betle extract were significantly lower (p < 0.05) than those of the controls. Results indicated that P. betle extract could increase the hydrophilicity of the SS samples; this could be due to the hydroxyl group and the polar bioactive compounds in P. betle extract. Greater hydrophilicity allows a sanitizer to spread on the surface more easily (greater wettability) [57]. When applied to biofilms, it could be deduced that P. betle extract may have increased the wettability of the biofilms, resulting in more efficient sanitization.

Conclusion
Overall in this study, the biofilm inactivation efficacy of the tested sanitizing agents against S. Typhimurium and L. innocua biofilms on pitted and smooth SS surfaces followed the trend of P. betle extract  BKC > NaClO > HP. The surface conditions of SS samples did not affect the biofilm inactivation efficacy of all tested sanitizers. P. betle extract increased the wettability of the pitted and smooth SS samples. This study indicates that P. betle extract may be an alternative to frequently used sanitizers for removing biofilms from food contact surfaces in food processing environments.